Most plant viruses are transmitted by insect vectors. Others are transmitted by mites, fungi or nematodes. Viruses that infect insects, such as baculoviruses, also enter the hemocoel of the insect from the gut through the action of virus coat proteins. Circulatively transmitted plant viruses enter the hemocoel (body cavity) of the insect vector. Viruses that infect insects also enter the hemocoel. In all cases, they have evolved specialized proteins to ensure their survival in the vector. This invention intends to exploit these viral proteins, and the genes encoding them, for presentation and delivery of toxins or other proteins to the vector or related organism. This may apply to non-vectors as well as vectors because, in some cases, viruses often interact with and enter the hemocoel of non-vectors. Furthermore, the viral proteins can be modified to interact with a wider range of organisms than occurs in nature. By facilitating efficient delivery of toxins, this invention can be used to genetically engineer pest-resistant plants, or to construct biopesticides.
Plant viruses can be transmitted in a circulative or noncirculative fashion. We define circulative transmission broadly as “any plant virus that must be actively transported across vector membranes and survive inside the vector to be transmitted” (Gray, S. M. [1996] Trends in Microbiology 4:259-264). By this definition, even some fungi can be considered circulatively transmitting vectors. In general, viruses are understood in the art as classifiable within groups which share common features of genetics, structure and the like. Examples of virus groups that are transported across insect vector membranes include: tospoviruses, plant reoviruses, plant rhabdoviruses, tenuiviruses, marafiviruses, luteoviruses, geminiviruses, enamoviruses, tymoviruses, comoviruses, and sobemoviruses (ibid: Gergerich, R. C. and Scott, H. A. [1991] Advances in Disease Vector Research. Harris, K. F., ed., pp. 114, Springer-Verlag). The fungally transmitted viruses include bymoviruses and furoviruses (op cit.; Jianping et al. [1991] Annals of Applied Biology 118:615-622). This invention encompasses all such viruses and their vectors and non-vectors which can acquire the virus.
The term “luteovirus” is used herein to encompass all viruses of the family Luteoviridae, including three major genera: Luteovirus, Polerovirus and Enamovirus. The RPV strain of Barley yellow dwarf virus (BYDV) described in examples herein, is also termed “cereal yellow dwarf virus—RPV” and classified in the genera Polerovirus, as the result of a recent change in nomenclature. However, since the majority of relevant art refers to BYDV, that nomenclature is retained herein.
Noncirculatively transmitted viruses “associate with the cuticular lining of the insect mouthparts or foregut and are released as the insect expels digestive secretions into the plant when it begins to feed. These viruses are not actively transported across vector-cell membranes, nor are they carried internally. The external cuticular lining of insects (and nematodes) extends well into the mouthparts and foregut, but is shed when the animal molts.” (Gray, S. M. [1996] Trends in Microbiology 4:259-264). Although the application herein exemplifies the use of plant virus proteins to deliver toxins to the hemocoel, noncirculatively transmitted viruses may be used similarly to present toxins to the surfaces of the gut or mouthparts. In principle, any virus which can enter the hemocoel of an insect from the gut through the action of viral protein can be employed, as described, to deliver a toxin to the insect.
The invention is exemplified herein by luteoviruses and their aphid vectors. They have the best-characterized circulatively-transmitted virus-vector interactions. However, the same principles can be applied for any virus-vector interaction and their exploitation to develop insect-resistant plants is included in this invention.
Luteoviruses can be transmitted only by aphids. The transmission mechanism is persistent and circulative. The virus enters the body cavity (hemocoel) of the aphid where it can remain for the life of the aphid. The aphid acquires virus by feeding on an infected plant. The virus particles (virions) are transported from the aphid hindgut into the hemocoel by a presumed receptor-mediated process across the hindgut epithelial cells. From the hemocoel, the virus is then transported across two more membrane barriers into the accessory salivary gland (ASG) (Gildow, F. E. et al. [1993] Phytopathology 83:1293-1302; Power, A. G. et al. [1995] in Barley Yellow Dwarf: 40 Years of Progress, St. Paul: APS Press, pp. 259-289), (FIGS. 1, 2). Subsequently, each time the aphid feeds, it transmits virus by secreting virus-laden saliva into the plant cells.
There is a high degree of vector specificity for different strains of luteoviruses (Power, A. G. et al. [1995] supra). However, many luteoviruses are transported across the hindgut membrane into the hemocoel, in many nonvector aphid species, (Gildow, F. E. et al. [1993] supra). Hence, vector specificity is thought to be enforced at the ASG barrier. Other viruses. such as potato leafroll virus, are transmitted across the midgut membrane. Nonluteoviruses are digested or excreted without uptake into the hemocoel. Thus, the hindgut epithelial receptors seem to be specific for luteoviruses (FIG. 2). This fact is exploited in the present invention.
The genome of luteoviruses consists of a single, positive sense, 5.7 kb RNA. The proteins needed for virus particles, aphid transmission, and virus movement within the plant all are expressed from a subgenomic RNA (sgRNA1) that is generated during virus infection but is not encapsidated in the virion (FIG. 3) (Miller, W. A. et al. [1997] Plant Disease 81:700-710). The luteovirus virion contains 180 copies of the coat protein (CP). About 5-10% of the CP subunits contain a long carboxy-terminal extension that protrudes from the virion (Filichkin, S .A. et al. [1994] Virology 205:290-299). This extension arises when ribosomes read through the stop codon of the CP open reading frame (ORF) during translation of sgRNA1 (FIG. 3), allowing translation of the downstream ORF5, resulting in a fused CP-readthrough domain (RTD) product (Brown, C. M. et al. [1996] J. Virol. 70:5884-5892).
The middle of the RTD (around amino acid 242) contains a labile peptide bond causing the C-terminal half to be cleaved in purified virus preparations (Filichkin et al. [1994] supra). Because purified virions are readily aphid transmitted, the C-terminal portion is unnecessary for aphid transmission. On the other hand, the N-terminal half of the RTD is required for aphid transmission (from aphid to plant) (Brault, J. et al. [1995] EMBO J. 14:650-659; van den Heuvel, J. F. J. M. et al. [1997] J. Virol. 71:7258-7265), but not for virion assembly or transport across the hindgut membrane into the hemocoel (Chay, C. A. et al. [1996] Virology 219:57-65).
The major CP itself is also required for transmission because intact virions are necessary to protect the viral RNA. In the hemocoel, the N-terminal half of RTD is bound by an abundant protein called symbionin which is produced by endosymbiotic bacteria (van den Heuvel et al. [1997] supra; Filichkin, S. A. et al. [1997] J. Virol. 71:569-579). This protein strongly resembles bacterial chaperonin groEL which ensures correct folding of many different proteins. However, the binding by symbionin does not resemble that of groEL with its substrate proteins (Hogenhout, S. A. et al. [1998] J. Virol. 72:348-365). The ability of symbionin to bind the luteovirus virion correlates with increased half-life of the virion in the hemolymph (van den Heuvel et al. [1997] supra).
It has been found that a sequence just 3′ of the CP ORF stop codon (proximal RT element) and a sequence located, surprisingly, 700-750 bases further downstream (distal RT element) are necessary for readthrough of the CP ORF stop codon during translation of sgRNA1 (Brown, C. M. et al. [1996] supra) (FIG. 3). The distal RT element still facilitates readthrough even after insertion of a reporter gene between it and the proximal element, causing the distal RT element to be located in the 3′ untranslated region (UTR) 2 kb downstream from the CP stop codon (Brown, C. M. et al. [1996] supra).
Current aphid control relies heavily on the use of chemical insecticides. Insecticide application to control transmission of viruses by aphids can have the unintended opposite effect of increasing virus spread by increasing plant-to-plant movement of aphids agitated by sublethal doses, and by killing aphid predators (Schepers, A. [1989] in Aphids: Their Biology, Natural Enemies and Control, Minks, A. D. et al. [eds.], Elsevier, Amsterdam, Vol C, pp. 123-139). Moreover, the efficacy of chemicals against aphids is limited because of the rapid evolution of insecticide-resistant aphids. Alternative, environmentally benign means of aphid control are required to maintain agricultural productivity.
Use of aphid-resistant crop cultivars has been effective for limiting aphid damage (Auclair, J. L. [1989] in Minks et al., supra, pp. 225-265; Thackray, D. J. et al. [1900] Ann. Appl. Biol. 116:573-582). Aphid-resistant maize lines (Walter, E. V. et al. [1946] J Am. Soc. Agron. 38:974-977) and wheat lines resistant to the Russian wheat aphid (Diuraphis noxia) (Quisenberry, S. S. et al. [1994] J. Econ. Entomol. 87:1761-1768) have been developed by conventional plant breeding techniques. Transgenic plants that resist insects have been constructed with agents that are active in the gut of insects. The most notable example is the use of the Bacillus thuringiensis toxin (Bt) genes, but no Bt toxin is known that affects aphids. Transgenic tobacco engineered to express a lectin was shown to confer protection against aphids (Hilder, V. A. et al. [1995] Transgenic Res. 4:18-25), and numerous transgenic plants have been produced that resist aphid-transmitted viruses by the use of virus-derived transgenes (Miller, W. A. et al. [1997] supra; Anon [1995] Genetic Engineering News 15:1; Wilson, T. M. A. [1993] Proc. Natl. Acad. Sci. 90:3134-3141).
A second approach toward insect pest control has been the use of baculoviruses, which are insect specific viruses. Some of these viruses have been used to deliver a variety of insect-specific toxins that, are active in the hemocoel but not in the gut of the insect. For example, recombinant baculoviruses have been developed for control of lepidopteran (moth) pest species (Bonning, B. C. et al. [1996] Annu. Rev. Entomol. 41:191-210). These baculoviruses have been engineered to produce insect hormones such as diuretic hormone (Maeda, S. [1989] Biochem. Biophys. Res. Comm. 165:1177-1183), enzymes such as juvenile hormone esterase (Bonning, B. C. et al. [1997] Proc. Natl. Acad. Sci. USA 94:6007-6012), and insect-specific toxins derived from venomous species such as scorpions and parasitic wasps (McCutchen, B. F. and Hammock, B. D. [1994] in Natural and Derived Pest Management Agents, Hedin, P. et al. [eds.], #551 ed., Washington, D.C.: Am. Chem. Soc., pp. 348-367. ACS Symposium Series; Hughes, P. R. et al. [1997] J. Invert. Pathol. 69:112-118; Lu, A. et al. [1996] in Biological control: theory and applications in 1996. 7:320; Gershburg, E. et al. [1998] FEBS Lett. 422:132-136; Jarvis, D. L. et al. [1996] in Biological control: theory and applications in 1996,7:228). These insecticidal proteins and peptides cannot be exploited for aphid control at present, because no system exists to deliver them into the hemocoel. Baculoviruses do not infect aphids, and although viruses such as Rhopalosiphum padi virus are known that infect aphids, there is insufficient knowledge of their biology and genetic structure for engineering for aphid control. Furthermore, the release of replicating, transgenic viruses poses greater risk to the ecosystem than expression of nonreplicating viral genes limited to the crop plant, as we disclose here. In the present invention, transgenically expressed, nonreplicating, plant viral structural proteins deliver insecticidal proteins or peptides into the hemolymph of aphids.
The toxin AaIT, that is derived from the venom of the North African scorpion Androctonus australis Hector, has a unique specificity for the nervous system of insects and some other arthropods such as crustaceans (Zlotkin, E. [1986] in Neuropharmacology and pesticide action, Chicester: Horwood; DeDianous S, et al. [1987] Toxicon 25:411-417). AaIT has no mammalian toxicity. Its strict selectivity for insects has been documented by toxicity assays, electrophysiological studies and binding assays (Zlotkin [1986] supra). AaIT is toxic to all insect species tested. These include over 15 species representing five orders of holo and hemimetabolous insects (Diptera, Coleoptera, Dictyoptera, Orthoptera and Lepidoptera) (Gershburg, E. et al. [1998] supra; Zlotkin [1986] supra; Herrmann, R. et al. [1990] Insect Biochemistry 20:625). The rapid excitatory paralysis induced by AaIT results from repetitive firing of the insect's motor nerves with resulting massive and uncoordinated stimulation of the respective skeletal muscles (Walther, C. et al. [1976] J. Insect Physiol. 22:1187-1194). The insecticidal activity of injected AaIT shows that this toxin is among the most potent toxic compounds to insects. AaIT has a molecular weight of only 8 kDa and has four disulfide bridges giving a highly organized conformation (Darbon, H. et al. [1982] Int. J. Peptide Protein Res. 20:320-332).
AaIT does not penetrate the lipophilic cuticle of insects (DeDianous, S. et al. [1988] Pestic. Sci. 23:35-40). To exploit this toxin for insect pest control purposes, various baculoviruses have been engineered to express AaIT (Gershburg, E. et al. [1998] supra; Jarvis, D. L. [1996] supra; Maeda, S. et al. [1991] Virology 184:777-780; McCutchen, B. F. et al. [1991] Bio/Technol. 91:848-852; Stewart, L. M. D. et al. [1991] Nature 352:85-88). These viruses infect the tissues of lepidopteran larvae and recombinant AaIT produced by the virus is exported from the virus-infected cell into the body cavity, resulting in paralysis of the insect. Expression of AaIT is highly effective for control of lepidopteran pest species and significantly reduces the amount of feeding damage caused. Because the nerves (the target site of AaIT) of Lepidoptera are unique in being covered by glial cells which act as a barrier to the toxin, AaIT is even more toxic to nonlepidopteran species. However, because baculoviruses do not infect aphids, they cannot be used for aphid control without modification. Also, because the site of action of the toxin is the nerves, transgenic plants expressing AaIT alone would not be aphid resistant because ingested toxin would pass through the insect.